Saponin and Fatty Acid Profiling of the Sea Cucumber Holothuria atra, α-Glucosidase Inhibitory Activity and the Identification of a Novel Triterpene Glycoside

Saponin-rich sea cucumber extracts have shown antidiabetic effects in a few reports. Although the triterpene glycosides of sea cucumbers are commonly isolated from their Cuvierian tubules, these are absent in Holothuria atra Jaeger. Therefore, this study intended to investigate the saponin profile in the body wall of H. atra, as well as to assess the α-glucosidase inhibitory activity of the H. atra extracts. The chemical profiling of sea cucumber extracts was conducted by UPLC-HRMS analysis. This resulted in the tentative identification of 11 compounds, 7 of which have not been reported in the H. Atra body wall before. Additionally, two triterpene glycosides were purified and their structures were elucidated based on HRMS and NMR data: desholothurin B (1), and a novel epimer, 12-epi-desholothurin B (2). Moreover, the fatty acid profile of the H. atra body wall was investigated by GC-MS. It was found that the Me90 fraction of the H. atra body wall showed the strongest α-glucosidase inhibitory activity (IC50 value 0.158 ± 0.002 mg/mL), thus making it more potent than acarbose (IC50 value 2.340 ± 0.044 mg/mL).


Introduction
Sea cucumbers (Holothuroidea) belong to the marine invertebrate Echinoderms. They are distributed in benthic areas, deep seas, and also in coral reefs. In traditional Chinese medicine, sea cucumbers are consumed for their beneficial health properties and are considered one of the delicacies of Asian-Pacific cuisine [1,2]. The traditional use of sea cucumbers may be linked to the presence of saponins, which were reported to possess antifungal [3,4], antiviral [5], antioxidant [6], and cytotoxicity activities [7][8][9][10].
Various species of sea cucumbers, including some Holothuria sp., have Cuvierian tubules, a gland expelled from the anus to defend the organism from marine animal attacks. The Cuvierian tubules contain a large variety of saponins, which are utilized as a chemical defense [11]. In addition, saponins can also be found in other parts of the sea cucumber, including the body wall and viscera [11,12], as well as in the seawater surrounding the sea cucumbers [13].
Thus, apart from higher plants [14], marine organisms such as sea cucumbers [15][16][17], sea stars [18][19][20][21], and sponges [22][23][24] can contain saponins. Saponins are glycosidic secondary Fresh H. atra sea cucumbers (12 kg) were collected in October 2016 from East Java, Indonesia. The fresh sea cucumbers were approximately 20 cm in length and 200 g in weight. Their identification was confirmed by the Research Center for Oceanography, Indonesian Institute of Sciences (voucher number B-173/IPK.2/IF/I/2017). The sea cucumbers were kept in a coolbox packed with ice. They were eviscerated to separate the body wall from their internal organs and washed with water. The body walls (3 kg) were dried in open air (±24 • C for 5 days, being exposed to direct sunlight for approximately eight hours a day), and were milled and sieved (sieve width 40 mm) to obtain a fine powder (600 g) using a disk mill machine. The moisture content of the dried H. atra was 9% and was determined using the drying method. The measurement was carried out by drying the samples in an oven at 105 • C until the constant weight was reached [52].

Preparation of Sea Cucumber H. atra Extract
The preparation of the sea cucumber extract was carried out according to Sottorf et al. (2013), with minor modifications [53]. An extract was prepared from the dried and milled material (500 g) using a soxhlet apparatus, and 6 L of CH 2 Cl 2 and 6.5 g of CH 2 Cl 2 extract (DCM) were resuspended in MeOH 90% (Me90), and then partitioned with 200 mL petroleum ether. Both subfractions were dried under reduced pressure at 40 • C. The residue after the initial soxhlet extraction was macerated with 13 L of MeOH 80%, and it was concentrated under reduced pressure and freeze-dried to obtain a methanolic crude extract (118 g). Figure 1 summarizes the general procedure of the extraction, fractionation, and chemical proflling of the sea cucumber H. atra. The UPLC-HRMS analysis of the H. atra body wall extracts led to the tentative identification of triterpene glycosides in ESImode ( Table 1). The molecular structure of triterpene glycoside identified from H. atra body wall is given in Table 2. The types of sapogenin, sapogenin side chains and also glycosidic moieties of triterpene glycosides from H. atra bodywall are shown in Figure 2. Atypical structure of sapogenin from triterpene glycoside identified from H. atra body wall including Calcigeroside B and Nobiliside II (=ananaside C) are shown in Figure 3 and Figure 4, respectively.

Fractionation of Triterpene Glycosides
The fractionation of the triterpene glycosides was carried out by means of a liquidliquid partition. In this procedure, the methanolic crude extract (60 g) was subjected to the liquid-liquid partition. It was solubilized in MeOH 50% (600 mL) and partitioned against n-hexane (600 mL). The n-hexane fraction and MeOH 50% fraction were concentrated under reduced pressure to obtain Hex fraction and Me fraction. The Me fraction was redissolved in 600 mL of water and was partitioned against 600 mL of n-BuOH. Both the Me fraction and the n-BuOH fraction (Bu fraction) were concentrated under reduced pressure and freezedried. The Me fraction and Bu fraction were submitted to UPLC-HRMS. The fractionation was carried out using open column chromatography and flash chromatography. The flash chromatography was performed on a Reveleris iES system from Grace (Columbia, MD, USA), with Reveleris ® Navigator™ software.
The subfraction MF/MG (2.3653 mg) was submitted to flash chromatography on a Reveleris C 18 cartridge by solid injection. The mobile phase used was H 2 O + 0.1% formic acid (A) and acetonitrile + 0.1% formic acid (B), with the following gradient: 0 min 95% for A and 5% of B retained for 15 min, which then changed to 50% for A and 50% for B at 52 min; from 97-111 min, a linear change to 0% for A and 100% for B, and from 111 min, a linear change to 95% for A and 5% for B, with a flow rate of 13 mL/min. The eluent was collected according to the signals measured by the evaporative light scattering detector Isolation was conducted on a semi-preparative HPLC-DAD-MS system (Waters, Millford, MA, USA) with Masslynx™software version 4.1. A Phenomenex Luna C18(2) 100 Å; 250 × 10.00 mm, 5 µm column was used together with a pre-column. The subfraction MFMG.9 (101.9 mg) was further purified by semi-preparative HPLC-DAD-MS with a C18 Luna column and the mobile phase H 2 O + 0.1% formic acid (A) and acetonitrile + 0.1% formic acid (B), and the following gradient: 0 to 5 min 42% of B, 30 min 50% of B, 35-40 min 100% of B, and 45-55 42% of B. The flow rate was 4.75 mL/min. The mass spectrometer was operated in ESI+ mode, with an MS scan range of m/z 250 to 800; V capillary 3.5 kV; V cone 50 V; V extractor 3V; V RF Lens 0.2 V; T source 125 • C; T desolvation 400 • C, and a desolvation gas flow of 750 L/h and a cone gas flow of 0 L/h to collect a compound with m/z 485.2 and m/z 763.3 (compound 1).

Structure Elucidation Using 1D and 2D NMR
A Bruker DRX-400 instrument (Rheinstetten, Germany) was used to record NMR spectra and was equipped with a 3 mm broadband inverse (BBI) probe or a 5 mm dual 1 H/ 13 C probe. Standard Bruker pulse sequences were used to record 1 H, 13 [54], using a XEVO-G2-XS QTOF mass spectrometer (Waters, Milford, MA, USA) coupled with an Acquity UPLC system. The system was operated with MassLynx 4.1 software (Waters, Milford, MA, USA). An HSS T3 RP18 column (1.8 µm, 2.1 × 100 mm) (Waters, Milford, MA, USA) was used to obtain separation. The following samples were analyzed using a UPLC Acquity system coupled with a Xevo G2-XS Q-Tof mass spectrometer (Waters, Milford, MA, USA), including subfractions MB, MC, ME, MF, and MG, and the Me fraction and Bu fraction. A total of two isolated compounds were also analyzed by UPLC-HRMS. The mobile phases used were (A) H 2 O + 0.1% FA and (B) ACN + 0.1% FA, and the gradient was set as follows: 3% of B (0-1 min), 100% of B (17-19 min), and 3% of B (21-25 min). The flow rate was 0.4 mL/min. The following settings were used for the mass spectrometer: a cone gas flow of 50 L/h; a desolvation gas flow of 1000 L/h; a source temperature of 120 • C; and desolvation at 550 • C. The samples were analyzed in MSe mode, thus obtaining information from the molecular ions and mass fragmentation data simultaneously. The MS data were recorded in ESI+ and ESI-mode with an MS scan range from m/z 50 to 1500.

GC-MS Analysis
The GC-MS analysis was conducted according to Dendooven et al. (2021), with minor modifications [55]. The petroleum ether (PE) fraction and MeOH 90% fraction were submitted to GC-MS analysis. The GC-MS analysis was carried out using a Trace GC Ultra (Thermo Fisher Scientific, Waltham, MA, USA), equipped with a capillary column (Restex Rxi5HT (30 m × 0.25 mm, 0.25 µm film thickness) (Chrom Tech, Apple Valley, MN, USA)). A 100 µL mix of fatty acid methyl esters containing 37 compounds was analyzed simultaneously, as is the reference standard. Helium (Praxair Technology, Danbury, CT, USA) was used as carrier gas with a flow rate of 0.1 mL/min. The splitless injection volume was 1 µL, and the inlet was heated to 250 • C. The oven temperature program was set as stated: 3 min of isothermal at 170 • C, increasing the temperature by 3 • C/min to 230 • C, and finishing with 15 min of isothermal at 230 • C. The MS analysis was performed with a DSQ mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) in scan mode within an m/z range of 50-1000 µ, with a run time of 41.75 min and scanning starting after 6 min. The GC-MS temperature of the ion source was 250 • C.

α-Glucosidase Inhibition Assay
The α-glucosidase inhibition was assessed using the method described by Su et al. (2013 andTrinh et al. (2016), with a slight modification [56,57]. In total, 100 µL of the αglucosidase enzyme solution from Saccharomyces cerevisiae (0.2 U/mL in a 0.1 M phosphate buffer at pH 6.8) and 50 µL of the sample dissolved in the phosphate buffer containing 6% DMSO 0.25-20 mg/mL), and were incubated in a transparent 96-well plate at 37 • C for 15 min. Next, 50 µL of PNPG (5 mM in a 0.1 m phosphate buffer) was added. The PNPG was hydrolyzed to release p-nitrophenyl (PNP), and this process was monitored at 405 nm for 30 min at 37 • C, with a BioTek Eon microplate reader (Winooski, VT, USA)) using the Gen5 version 2.06 software. The blank sample was treated in a similar way, but contained a 0.1 M phosphate buffer at pH 6.8 (containing DMSO 6%) instead of a test compound. Acarbose (≥95%, Sigma Aldrich) was used as a positive control, and all the measurements were repeated three times. The IC 50 values were calculated with GraphPad Prism 6 software (GraphPad Software Inc, La Jolla, CA, USA). The IC 50 values were subjected to one-way analysis of variance (ANOVA) and the Tukey post hoc test, using GraphPad Prism 6 software to assess any significant differences among the treatments. P-values < 0.05 were considered significant.

Tentative Identification of Triterpene Glycosides from H. atra Body Wall
The UPLC-HRMS analysis of the H. atra body wall extracts led to the tentative identification of triterpene glycosides in ESImode ( Table 1). The molecular structure of triterpene glycoside identified from H. atra body wall is given in Table 2. The types of sapogenin, sapogenin side chains and also glycosidic moieties of triterpene glycosides from H. atra bodywall are shown in Figure 2. Atypical structure of sapogenin from triterpene glycoside identified from H. atra body wall including Calcigeroside B and Nobiliside II (=ananaside C) are shown in Figures 3 and 4, respectively.

Purified Saponins from H. atra Body Wall
A total of two compounds were isolated from the combined MF/MG subfraction. These two subfractions were combined because they showed a similar pattern after NP-TLC analysis. Subsequently, the combined fraction was submitted to RP flash chromatography, and the fractions with similar NP-TLC profiles were again combined, resulting in 15 subfractions. Subfractions 8 and 9 were selected for the semi-preparative HPLC-DAD-MS, which led to the isolation of two isomeric compounds.
The structure elucidation of compound 1 was performed based on 1D and 2D-NMR analysis (spectra are provided as supplementary material, Figures S1-S8), as well as on UPLC-HRMS measurement (mass spectrum provided as supplementary material, Figure  S14), leading to its identification as desholothurin B (desulfated holothurin B) ( Figure 5, Table 3). Indeed, the 13 C chemical shifts of compound 1 were in agreement with those previously reported [84,85] for the desulfated derivative of holothurin B, which was derived from holothurin B by means of refluxing in dioxane/pyridine (1:1). Kobayashi and co-workers obtained the desulfated holothurin B from holothurin B in a similar way [11]. Thus, in these studies, desulfated holothurin B was not obtained as natural product but as derivative of holothurin B. However, in 1987, Oleinikova and Kuznetsova had already studied the glycosidic fraction of H. atra and reported that this fraction contained 2.15% of desulfated holothurin B, while holothurin B was found to be the major constituent (84.2%) [86]. Unfortunately, no experimental data regarding this identification were provided, but nevertheless, this publication must be considered the first and only publication thus far to report the presence of desholothurin B as natural product. In addition, a second compound was isolated, which showed the same m/z-value in UPLC-HRMS, but a different retention time.  Figures S14-S17). Furthermore, also noteworthy was a difference in solubility that was noticed when preparing the compounds for NMR analysis: while compound 1 was soluble in methanol-d 4 , this was not the case for compound 2, and the latter was therefore analyzed in pyridine-d 5 . Given the low amount of sample available, unfortunately no decent 13 C-and DEPT-spectra could be recorded for compound 2. Nevertheless, a complete assignment could be performed based on the 1 H-and 2D-spectra (supplementary material, Figures S9-S13), with the exception of C-18.
TLC analysis. Subsequently, the combined fraction was submitted to RP flash chromatog raphy, and the fractions with similar NP-TLC profiles were again combined, resulting in 15 subfractions. Subfractions 8 and 9 were selected for the semi-preparative HPLC-DAD MS, which led to the isolation of two isomeric compounds.
The structure elucidation of compound 1 was performed based on 1D and 2D-NMR analysis (spectra are provided as supplementary material, Figures S1-S8), as well as on UPLC-HRMS measurement (mass spectrum provided as supplementary material, Figur S14), leading to its identification as desholothurin B (desulfated holothurin B) (Figure 5  Table 3). Indeed, the 13 C chemical shifts of compound 1 were in agreement with thos previously reported [84,85] for the desulfated derivative of holothurin B, which was de rived from holothurin B by means of refluxing in dioxane/pyridine (1:1). Kobayashi and co-workers obtained the desulfated holothurin B from holothurin B in a similar way [11] Thus, in these studies, desulfated holothurin B was not obtained as natural product bu as derivative of holothurin B. However, in 1987, Oleinikova and Kuznetsova had already studied the glycosidic fraction of H. atra and reported that this fraction contained 2.15% of desulfated holothurin B, while holothurin B was found to be the major constituen (84.2%) [86]. Unfortunately, no experimental data regarding this identification were pro vided, but nevertheless, this publication must be considered the first and only publication thus far to report the presence of desholothurin B as natural product. In addition, a second compound was isolated, which showed the same m/z-value in UPLC-HRMS, but a differ  Figures S14-S17). Furthermore, also notewor thy was a difference in solubility that was noticed when preparing the compounds fo NMR analysis: while compound 1 was soluble in methanol-d4, this was not the case fo compound 2, and the latter was therefore analyzed in pyridine-d5. Given the low amoun of sample available, unfortunately no decent 13 C-and DEPT-spectra could be recorded fo compound 2. Nevertheless, a complete assignment could be performed based on the 1 H and 2D-spectra (supplementary material, Figures S9-S13), with the exception of C-18.   The majority of 13 C-NMR signals observed for compound 2 were similar to those of compound 1 (Table 3), and could be assigned by comparison with the NMR assignment of compound 1, in combination with the interpretation of the 2D-NMR spectra. However, six out of thirty of the 13 C chemical shifts showed a difference >2 ppm compared to the signals of 1, while the other signals typically differed less than 1 ppm, again indicating a structural difference between the two compounds. These 13 C-NMR signals were assigned to C-9 and C-11-C-14 of ring C, as well as C-17 of the holostane sapogenin moiety as follows: firstly, a signal at 151.2 ppm showed a cross peak in the HMBC-spectrum with the 1 H-signal of the methylgroup in postion 19, and was assigned to C-9, since all other signals in close proximity to C-19 were already assigned. A total of two rather downfield 1 H-signals, found at 5.62 and 5.25 ppm, showed a COSY interaction and were found to correspond to positions 11 and 12. Their exact positions were deduced after the assignment of the 13 C-signals of C-11 and C-12: the 13 C-signal at 119.5 ppm complies with a methine substructure, as in position 11, while the 13 C-signal at 66.9 ppm complies with a hydroxylated tertiary carbon. Therefore, based on the HSQC-spectrum, the signals at 5.62 and 5.25 ppm were assigned to H-9 and H-11, respectively. The two quaternary carbon signals, C-13 and C-14, at 65.3 and 48.7 ppm, respectively, both showed a cross peak in the HMBC-spectrum, with the 1 H-signal of the methylgroup in position 30 (1.37 ppm), thus supporting their presence in these positions. Given the proximity of the hydroxyl group in position 12, and the carbonylgroup in position 18, C-13 was assigned the more downfield signal. A 13 C-signal, differing from the signals observed for compound 1, remained: a quaternary carbon at 87.4 ppm. This signal was assigned to position C-17, which was supported by an HMBC-correlation with the 1 H-signal of the methyl group in position 21 (2.04 ppm).
The highest discrepancy of the chemical shifts observed for the two compounds was found between the signals assigned to C-12 and C-13 (5.6 ppm), and it was inferred that the structural difference of compound 2 and compound 1 occurs in this part of the molecule. In fact, the only plausible explanation for these observed differences is that compound 2 is an epimer of compound 1, bearing the 12-hydroxyl group in a different configuration. Previously, several cases were reported in which the relative configuration of the 12-OH group of a Holothuria triterpene glycoside was determined based on NOESY correlations [3,7,8]. It was stated that, in the case of an α-configuration, a correlation of H-12 with a proton signal originating from the methyl group attached to C-20 (H 3 -21) was observed [3,7,8,84], and in general, this relative configuration is reported for this type of compound. When comparing the NOESY spectra obtained for compounds 1 and 2 (supplementary Figures S5 and S13), in the first case, a correlation was indeed observed between H-12 (4.53 ppm) and H 3 -21 (1.50 ppm), while no correlation was observed for the H-12 (5.31 ppm) and H 3 -21 (1.21 ppm) of compound 2. Thus, these results are in line with the proposed structure of compound 2, bearing a 12β-OH group, i.e., 12-epi-desholothurin B ( Figure 5). To the best of our knowledge, this compound has not been reported before.
With regards to the other stereocenters, the same relative configurations are assigned to compounds 1 and 2, as previously reported for holothurin B. This configuration is most commonly reported for the triterpene glycosides of the Holothuria genus in general [73,84,85].

Fatty Acid Profile of the H. atra Body Wall
The Me90 was analyzed by GC-MS, and this led to the identification of 11 fatty acids, including both saturated and unsaturated fatty acids (Table 4). Several fatty acids were previously reported in H. atra [36,46,87], while two of the identified fatty acids are reported herewith for the first time in H. atra.

α-Glucosidase Inhibition Assay
The α-glucosidase inhibition assay was conducted to determine the anti-diabetic potential of the H. atra body wall. Acarbose was used as a positive control (Table 5). In order to further study the activity of the Me90 fraction, commercially available fatty acids were also tested against α-glucosidase (Table 6). A previous study, conducted by Su and team, revealed that fatty acids are potent α-glucosidase inhibitors [43,44,56]. The IC 50 values of the MG subfraction and the Me90 fraction did not differ significantly. The IC 50 values of fatty acids (palmitoleic acid, arachidonic acid, and eicosapentaenoic acid) were significantly different to those of the acarbose.

Discussion
In this study, 11 triterpene glycoside compounds were tentatively identified in the sea cucumber H. atra. While the applied UPLC-HRMS analysis can provide information on the molecular weight of the saponin, and often of the sapogenin, a distinction between the triterpene glycosides with isomeric sapogenin moieties and those with glycosidic moieties is, in many cases, impossible. Therefore, for several of the identified signals, more than one possible identified compound was listed in Table 1.
In general, triterpene glycosides contain an aglycone which is either a 3β-hydroxyholost-7-ene or a 3β-hydroxyholost-9(11)-ene aglycone [88]. Holothuria triterpene glycosides contain an aglycone moiety with a holostanol skeleton. Puspitasari and co-workers grouped the sapogenins of the Holothuria triterpene glycoside into six types [32]. Such sapogenins are common sapogenins in the genus Holothuria [32,80,89]. Triterpene glycosides are also classified based on their types of side chain, including those with a tetrahydrofuran group and those with a linear side chain. With regards to the side chain, 25 types are commonly identified within the Holothuria genus. In this work, 11 types of triterpene glycosides were identified from the body walls of H. atra [32,89].
In this study, only one compound, calcigeroside B (Figure 3), contains a double bond between C-7 and C-8, while a 9(11)-double bond is present in all the other triterpenoid glycosides of H. atra. Despite calcigeroside B, nobiliside (II) or ananaside C (Figure 4) also has an atypical structure of triterpene glycosides, and this compound was previously identified from H. nobilis [90].
With respect to the nomenclature of the sea cucumber saponins, it is important to be aware of the fact that, in some cases, the same compound was named differently by different authors, while, in other cases, different compounds got the same name. Honey-Escandon stated that the chemical nomenclature of the triterpene glycosides could be inconsistent and homonyms occur [89]. In this case, holothurin A5 was identified in H. atra with the molecular formula C 54 H 83 O 28 SNa and a nominal mass of 1234 [37], while it was reported later with the same compound name, holothurin A5, but with the different molecular formula C 54 H 85 O 28 SNa and a nominal mass of 1236 [66]. Wu et al. reported ananaside C as a new compound from Thelenota ananas in 2007. However, a triterpene glycoside with the same formula was reported as nobiliside II from H. nobilis by Zhang in 2011 [89,90].
In this study, the sea cucumber saponins identified by means of UPLC-HRMS analysis were tentatively identified as sulfated saponins. In contrast, the two compounds isolated from the body wall of the H. atra: desholothurin B (1) and epi-desholothurin B (2), were not sulfated [84]. Van Dyck et al. [12] and Omran et al. [36] stated that H. atra contains only sulfated triterpene glycosides, such as echinoside A, echinoside B, and holothurins B/B4, B1, B2, and B3. However, according to Oleinikova and co-workers, desulfated holothurin B was detected in H. atra [86].
According to Ridzwan and co-workers, the body wall of the H. atra contained 57.04% of saturated fatty acids, 4.31% of monounsaturated fatty acids, and 38.64% of polyunsaturated fatty acids [46]. With regards to the lipid fraction of the H. atra, the saturated fatty acids were myristic acid, pentadecanoic acid, palmitic acid, stearic acid, arachidic acid, heneicosanoic acid, and behenic acid, while the other identified fatty acids were unsaturated. Myristic acid, palmitoleic acid, palmitic acid, stearic acid, arachidonic acid, eicosapentanoic acid, arachidic acid, behenic acid or docosanoic acid, and nervonic acid were previously reported [36,46], while nervonic acid and heneicosanoic acid are reported in H. atra for the first time. Arachidic acid was previously identified in H. edulis and Bohadschia marmorata. Nervonic acid was reported in H. polii, H. edulis, and B. marmorata [36].
As for the α-glucosidase inhibition, the MG subfraction was found to be more active than acarbose. The following compounds were tentatively identified in subfraction G: leucospilatoside A, holothurin B3 or 24-dehydroechinoside B, echinoside A (=holothurin A2), and 17-dehydroxyholothurin A (=fuscocineroside C), scabraside A, 24-dehydroechinoside A, or fuscocineroside B. Unfortunately, the amount of isolated saponins was not sufficient enough to allow for individual testing in the α-glucosidase inhibition assay, and therefore, the hypothesis that the isolated saponins present in the MG subfraction contributed to the observed inhibitory activity could not be confirmed.
The inhibition of α-glucosidase activity in the small intestine is one strategy to suppress blood glucose levels. α-glucosidase, located on the brush-border of the small intestine, plays a pivotal role in carbohydrate digestion by hydrolyzing starch into monosaccharides (glucose). By inhibiting the α-glucosidase activity, carbohydrate digestion and glucose production can be delayed. Therefore, less glucose will be absorbed into the blood stream, which can reduce hyperglycemia [47]. This study revealed that some lipophilic and hydrophilic fractions of H. atra body walls can inhibit α-glucosidase. These fractions were found to contain several fatty acids and triterpene glycosides. Further studies will need to be conducted to determine the specific inhibitory activity of these compounds and to assess whether H. atra could play a role in the management of glucose levels in diabetic patients.

Conclusions
In conclusion, the phytochemical profile of H. atra body walls was investigated and 13 triterpene glycosides and 11 fatty acids were (tentatively) identified. The saponin 12-epidesholothurin B (2) was purified and identified for the first time. It was found that several fractions of H. atra body walls showed α-glucosidase inhibitory activity.